Protocol to establish a genetically engineered mouse model of IDH1-mutant astrocytoma

Summary Lower-grade gliomas exhibit a high prevalence of isocitrate dehydrogenase 1 (IDH1) mutations, but faithful models for studying these tumors are lacking. Here, we present a protocol to establish a genetically engineered mouse (GEM) model of grade 3 astrocytoma driven by the Idh1R132H oncogene. We describe steps for breeding compound transgenic mice and intracranially delivering adeno-associated virus particles, followed by post-surgical surveillance via magnetic resonance imaging. This protocol enables the generation and use of a GEM to study lower-grade IDH-mutant gliomas. For complete details on the use and execution of this protocol, please refer to Shi et al. (2022).1


SUMMARY
Lower-grade gliomas exhibit a high prevalence of isocitrate dehydrogenase 1 (IDH1) mutations, but faithful models for studying these tumors are lacking. Here, we present a protocol to establish a genetically engineered mouse (GEM) model of grade 3 astrocytoma driven by the Idh1 R132H oncogene. We describe steps for breeding compound transgenic mice and intracranially delivering adeno-associated virus particles, followed by post-surgical surveillance via magnetic resonance imaging. This protocol enables the generation and use of a GEM to study lower-grade IDH-mutant gliomas. For complete details on the use and execution of this protocol, please refer to Shi et al. (2022). 1

BEFORE YOU BEGIN
This protocol describes the specific steps required to generate a genetically engineered mouse (GEM) model of grade 3 IDH-mutant astrocytoma. The tumors arising in this mouse model recapitulate key histologic and lineage features of grade 3 IDH-mutant human astrocytomas and are driven by mutant IDH. Our approach entails two primary activities: 1) breeding compound transgenic mice, and 2) delivering AAV particles into the brains of compound transgenic mice via stereotactic intracranial injections. In addition to describing how to establish this IDH-mutant glioma GEM model, this protocol also covers establishment of isogenic control GEM models in which glioma formation is infrequently observed. This protocol can also be used to produce glioma stem-like cell (GSC) lines and allograft models from autochthonous tumors that form in the IDH-mutant glioma GEM model. All of these approaches are described in recently published work. 1 The IDH-mutant glioma GEM model described in this protocol features oncogenic Pik3ca H1047R and Idh1 R132H transgenes and a Cas9 transgene. For simplicity, we refer to the triple transgenic mice as ''PIC'' mice. In addition to PIC mice, this protocol can also be used to generate mice with Idh1 R132H and Cas9 alleles (IC), Pik3ca H10474 and Cas9 alleles (PC), and a Cas9 allele alone (C). Naïve mice are phenotypically wild-type, as all three genes are constitutively repressed by lox-STOP-lox (LSL) elements. Intracranial injection of AAV particles harboring a Cre recombinase cDNA and sgRNAs b. To make additional isogenic control GEM models lacking the Pik3ca H1047R transgene, cross H11 LSL-Cas9+/+ ;Idh1 tm1Mak/WT mice with wild-type FVB mice. This breeding produces IC (H11 LSL-Cas9+/À ;Idh1 tm1Mak/WT ) and C (H11 LSL-Cas9+/À ;Idh1 WT/WT ) mouse strains. Verify genotype with progeny as above in step 2. c. Genotype the R26 locus using the Pik3ca-1 Tg and ROSA WT Transnetyx probes.
Note: Stereotactic intracranial AAV injections, described later in this protocol, should be performed on male and female mice at least 6 weeks and no more than 7 months old. Use of younger mice may negatively affect reproducibility of injections and subsequent tumor formation due to changes in brain architecture during development.

Preparation of AAV
Timing: 2-4 weeks 5. Propagate pAAV plasmid pAAV2-sgTrp53-sgAtrx-EFS-Cre that can be used to produce AAV. 6. High-titer AAV preparations can be produced through commercial sources or in individual labs.
We have used Charles River (previously Vigene Biosciences) for AAV production. Viral titers should be in the range of 1 3 10 13 -1 3 10 14 genome copies/mL.

CRITICAL:
The titer of AAV preparations should be in the 1 3 10 13 -1 3 10 14 genome copies/mL range. The volume that can be injected intracranially is limited to approximately %3 mL, and high-titer virus is thus needed to ensure adequate infection rates of Figure 1. Breeding scheme Breedings required to generate PIC mice for intracranial injection of AAV. Note that this breeding scheme also generates PC mice, which can be used to generate an isogenic IDH1-wild-type control if desired. Breedings to generate IC and C mice (bottom right) are optional though can be used for additional controls. Also note that some of the depicted matings may generate multiple genotypes not shown, and the progeny of interest should be determined through the genotyping steps outlined in ''generation of compound transgenic mouse strains.'' neural stem and progenitor cells in the subventricular zone, the presumed cell of origin for autochthonous gliomas in this GEM model.

Preparation of materials used for surgery
Timing: 1 h 7. Sterilize all surgical tools. a. Place surgical scissors, drill bits, syringe/needles, forceps, stapler, and staples in autoclavable container. b. Use a gravity autoclave setting at 121 C for a minimum of 30 min. c. Remove tools and let cool to room temperature (20 C-23 C). 8. Prepare aliquots of sterile PBS.
a. Prepare sterile 1.5 mL Eppendorf tubes with 1 mL aliquots of sterile PBS inside a laminar flow hood. These will be used for flushing the syringe/needle used for intracranial injection between injections. Prepare at least one aliquoted tube per mouse to be injected. b. Separately, prepare a 50 mL tube of sterile PBS inside a laminar flow hood. This will be used for rinsing surgical tools (scissors, forceps) between mice. 9. Prepare aliquots of iodopovidone-based disinfectant.
a. Prepare 1 mL aliquots of iodopovidone disinfectant solution (such as Wescodyneâ) in sterile Eppendorf tubes. These will also be used to clean the syringe/needle used for intracranial injection between injections. Prepare at least one aliquoted tube per mouse to be injected. b. Separately, prepare a 50 mL tube of iodopovidone disinfectant solution. This will be used for disinfecting surgical tools (scissors, forceps) between mice. 10. Aliquot AAV (1 mL per mouse) into a sterile Eppendorf tube. Store AAV on ice throughout surgeries. AAV should be stored as single-use aliquots at À80 C long-term and should not undergo multiple freeze-thaw cycles.
Pause point: Protocol can be paused here for days to weeks before day of surgery.

Setup of anesthesia, surgery, and recovery areas
Timing: 30 min For the subsequent steps, use appropriate personal protective equipment, including sterile surgical gloves, surgical mask, surgical cap or hair net, and lab coat or disposable gown.
11. Prepare the laminar flow hood to perform intracranial injections (Figure 2, top). a. Sanitize the surfaces of the hood with a suitable disinfectant. b. Plug in and power on the stereotactic frame and the control panel. c. Remove the surgical tools (forceps, scissors, stapler, and staples) from the box used to autoclave. Lay tools out on sterile gauze in the hood. d. Assemble a guide needle (3 mL syringe with a 25 G needle attached) and lay it on sterile gauze in the hood. 12. Prepare separate, adjacent hood for anesthesia/analgesia administration ( Figure 2, bottom left).
a. Use appropriate personal protective equipment. b. Sanitize the surfaces of the hood with a suitable disinfectant. c. Prepare an empty cage to be used for holding mice immediately prior to ketamine/xylazine anesthesia administration. d. Prepare sterile cotton-tipped applicators and ophthalmic ointment. e. Set up a scale and box that can be used to weigh mice. 13. Prepare cages for surgery/anesthesia recovery (Figure 2, bottom right).

OPEN ACCESS
a. Set up a slide warmer set to 35 C-38 C with 1-2 empty cages to be used for immediate postsurgical recovery. b. Set up a clean cage with food and antibiotic-containing water (ex. enrofloxacin). Post-surgical mice should be given antibiotic-containing drinking water for 7 days postoperatively for infection prophylaxis.
Note: The above setup designates one hood for surgery preparation and anesthesia administration and a second, adjacent hood for intracranial injections. Alternative hood setups such as a single hood may be considered, for example if using inhaled anesthetics (e.g., isoflurane) to anesthetize mice.

STEP-BY-STEP METHOD DETAILS
The protocols below detail the component steps for stereotactic intracranial injections of AAV. Variations in the steps are noted when injecting cells to generate GEM-derived allografts (see generation of GEM-derived allografts).

Anesthetizing the mouse
Timing: 10-30 min The following steps detail the procedures for administering injectable ketamine/xylazine anesthetic and for assessing depth of anesthesia.
1. Prepare the ketamine/xylazine anesthetic solution a. Make the ketamine/xylazine solution as described in the materials and equipment section of this protocol. b. Filter sterilize solution by passing the solution through a 0.22 mm filter with a syringe. c. Weigh mouse to be anesthetized. Record weight (in grams) and return mouse to cage. d. Use a 1 mL syringe to slowly draw up volume of ketamine/xylazine mixture at 5 mL per gram of body weight, achieving a final dose of 140 mg/kg ketamine and 12 mg/kg xylazine. e. Turn the syringe needle up and plunge the syringe to expel any air bubbles in the syringe. f. Set syringe down on a clean surface. Use caution with the exposed needle tip. 2. Administer the ketamine/xylazine solution a. Hold and stabilize the mouse in one hand with a controlled and firm grip. b. Use an alcohol pad to sterilize the intraperitoneal injection site. c. Anesthetize the mouse by injecting it with the pre-filled 1 mL syringe intraperitoneally. d. Place mouse in separate clean cage to allow for anesthesia to take effect. 3. Assess depth of anesthesia a. The mouse should exhibit slowed movements within 1-2 min of injection. b. Begin checking for depth of anesthesia approximately 5-10 min following injection. Perform toe pinch test and assess for presence of reflexive movements after a firm pinch to one of the hind feet of the mouse. Adequate depth of anesthesia may take 15 min or longer following injection. 4. Once mouse is fully anesthetized, apply ophthalmic ointment to both eyes (Figure 3, right) and use clippers to shave the fur above the skull. Duration of anesthesia is approximately 1-2 h.
CRITICAL: Ensure that the mouse is fully anesthetized and does not move in response to the toe pinch test. It is important to anesthetize the mouse fully prior to performing intracranial injections. The skull is well-innervated, and this procedure can cause pain if the mouse is not fully anesthetized. Intracranial surgeries performed without sufficient anesthesia can cause movement during skull drilling and undue pain and suffering to the mouse.
Note: We utilize ketamine/xylazine, extended-release buprenorphine, and meloxicam for anesthesia/analgesia for these surgeries, although alternative anesthetics may be used (e.g., isoflurane).
5. Administer local anesthetic a. Using previously recorded weight of mouse, draw up appropriate volume of 0.5 mg/mL meloxicam in 1 mL syringe to achieve 0.5 mg/kg dose. b. Turn syringe needle up and expel any bubbles in syringe. c. Administer meloxicam subcutaneously to the scruff between the ears (Figure 3, left). CRITICAL: Mouse positioning should be very stable prior to proceeding to next step. If the mouse is not securely positioned into the stereotactic frame, the mouse may move during skull drilling, leading to inaccurate burr hole creation.
7. Use an alcohol pad to disinfect the skin surface overlaying the skull. 8. Make a 1-1.5 cm incision in the craniocaudal direction to expose the skull ( Figure 5). 9. Use a cotton-tipped applicator and gentle pressure to rub the outer membrane away from the exposed skull ( Figure 5, top left).
Note: The membrane is clear and will feel slick. The underlying skull will feel noticeably less slick and rougher when rubbed with the cotton-tipped applicator.
10. Position the guide needle (3 mL syringe with a 25 G needle attached) into the syringe holder on the stereotactic frame ( Figure 5, top right).
Note: The guide needle is used to zero the coordinates on the stereotactic frame. The coordinates from this positioning will be used to determine the site of virus injection.
11. Position the needle directly centered on the bregma ( Figure 5, top right).

OPEN ACCESS
Note: A detailed image of the exposed skull anatomy is included in Figure 5 (top middle) to guide visualization of the anatomical landmarks. The guide needle tip may gently touch the skull to confirm accurate positioning, but do not pierce the skull with the guide needle.
12. Zero the anterior/posterior and medial/lateral coordinates on the stereotactic frame ( Figure 5, bottom).
CRITICAL: Ensure that the guide needle is positioned precisely at the location of the bregma. Inaccurate zeroing of the coordinates at the bregma may lead to inaccurate viral injection site.
13. With the guide needle still in place, use the stereotactic frame adjustments to move the guide needle 1 mm posterior and 1 mm lateral to the bregma. 14. Drill a burr hole into the skull at this coordinate position.
a. Position the tip of the guide needle 0.5-1 cm above the surface of the skull to indicate injection site. b. Use the handheld drill to carefully drill a small burr hole at this location on the skull (1 mm poster and 1 mm lateral to the bregma) ( Figure 6, left, middle). See troubleshooting 2. c. Apply gentle pressure and retract drill bit as soon as pressure abates, indicating successful passage of the drill bit through the skull. 15. Using a 5 mL syringe with affixed 1.5 in, 32 G needle, carefully draw up 1 mL AAV virus. Carefully inspect the syringe to confirm absence of bubbles.
CRITICAL: Ensure that there are no bubbles present in the syringe before proceeding to next step (see troubleshooting 3).
Note: Different injection coordinates are used when establishing GEM-derived allografts (3 mm anterior and 2 mm lateral to the lambda, 2.5 mm depth). See generation of GEMderived allografts. A different volume of cells may be used to inject a total of 1 3 10 5 cells, though this volume should not exceed 3 mL.
16. Remove the guide needle from the stereotactic frame and replace it with the 5 mL syringe/needle filled with 1 mL virus. 17. Position the syringe such that the needle is directly overlaying the hole drilled into the skull (Figure 6, right). 18. Lower the syringe/needle using the adjustments on the stereotactic frame such that the needle tip abuts the surface of the brain. Zero the dorsal/ventral coordinates on the stereotactic frame. 19. Lower the needle to a depth of 2.1 mm below the brain surface. 20. Slowly inject the virus into the brain (1 mL of virus over 3 min). Wait 1 additional minute after fully dispensing AAV particles to allow liquid to disperse in tissue before retracting needle.
CRITICAL: Injecting the virus slowly is important for minimizing the volume of AAV that leaks out of the brain. Excess leakage may contribute to formation of needle-track skullbased tumors.
21. Slowly remove the needle from the brain by manually using the dorsal/ventral controls of the stereotactic frame over a duration of approximately 60 s. Once the needle tip is visible outside of the brain and skull, the needle and syringe can be removed from the frame. 22. Close the incision site with a wound stapler (Figure 7, left). Use forceps to bring together the skin surface in order to approximate the skin prior to applying staples.
Note: Do not apply saline or other lubricants to the skull surface, as this may cause diffusion and leakage of the injected virus throughout the skull surface.
23. Place the mouse in the recovery cage on slide warmer (Figure 7, right). 24. As the mouse begins to wake from anesthesia and spontaneously move, draw up appropriate volume of buprenorphine in a 1 mL syringe to inject the mouse with 0.1 mg/kg subcutaneously.
Note: If using Ethiqaä (extended release buprenorphine), it is recommended to use a 20 G or 23 G needle, as the suspension is viscous.
25. Sterilize buprenorphine injection site with an alcohol pad and inject buprenorphine subcutaneously. 26. Place mouse in clean cage with an antibiotic-containing water (ex. enrofloxacin). Post-surgical mice should be given antibiotic-containing drinking water for 7 days postoperatively for infection prophylaxis. Staples should be removed 7-14 days after intracranial surgery.

Cleaning surgical instruments and inactivating leftover AAV particles
Timing: 5 min These steps outline the best practices for disinfecting surgical tools between each mouse operation, as well as long-term maintenance of the 5 mL syringe to prevent clogs.

Surgical instruments
27. Rinse all surgical tools (scissors, forceps, drill bit) with iodopovidone disinfectant, followed by sterile PBS. 28. Use sterile gauze to dry them.

Intracranial injection syringe
29. Dip the needle tip of the 5 mL syringe into an aliquot of povidone-iodine disinfectant, followed by sterile PBS. 30. Dry needle tip thoroughly with sterile gauze. Use caution to not bend the needle tip. 31. Draw up and expel sterile PBS 3-5 times to flush contents of syringe.
Note: If syringe does not draw up or draws up with bubbles, remove the plunger, back fill syringe with sterile PBS, and replace plunger.
Note: To optimally preserve use of the 5 mL syringe, use cleaning concentrate solution (see key resources table) per package instructions after concluding intracranial injections.
Inactivate AAV particles 32. Expose all solutions containing AAV particles and all solutions that have come in contact with AAV particles to 10% bleach for >15 min to inactivate virus. After inactivation, solutions can be discarded via sink disposal.

Post-surgical MRI surveillance
Timing: 8-10 months This section describes steps necessary for MRI surveillance of mouse brains, which is used to assess for intracranial tumor growth.

OPEN ACCESS
osteosarcomas and protracted tumor latency associated with the GEM model. 1 The procedures for anesthesia and intracranial injection to produce orthotopic allografts are largely identical to the above outlined steps to generate the GEM. The critical points of difference are highlighted in the above protocol steps and emphasized below.
34. PIC tumor-derived GSC line creation a. After brain tumor formation on MRI is observed, euthanize the mouse. b. Immediately harvest the brain and isolate tumor-containing tissue by gross inspection. c. Dissociate tumor tissue into a single cell suspension using the Miltenyi Neural Tissue Dissociation Kit (see key resources table). d. Culture cells in 5% CO 2 and 5% O 2 on ultra-low adherence plates to select for neurosphereforming GSCs.
CRITICAL: GEM-derived neurospheres must be cultured at 5% O 2 . Attempts to establish murine neurosphere cell lines from this GEM model under ambient oxygen conditions were unsuccessful.
35. Intracranial injection a. Inject 1 3 10 5 cells (prepared in phosphate-buffered saline) into female ICR SCID mice. Note that the preparation of cells should be at a concentration of at least 3.3 3 10 4 cells/mL so as not to exceed 3 mL injection volume.
CRITICAL: Use of injection volume R3 mL and/or excessively rapid injections may result in leakage of cell suspension out of the brain and affect glioma latency and promote the development of ectopic tumors (see problem 5).
b. Inject cells at 3 mm anterior, 2 mm lateral to the lambda ( Figure 5) and 2.5 mm below the brain surface. 36. Monitor tumors via MRI a. Monitor mice for tumor formation with serial MRI scans as in step 33.

EXPECTED OUTCOMES
Visible tumors are present on MRI scans starting as early as 8 months after AAV injection in PIC mice ( Figure 8). DF-AA27 cells derived from a glioma arising in a PIC mouse 1 formed allografts visualizable on MRI scans 1-4 months after intracranial implantation (Figure 8).

LIMITATIONS
The latency period between AAV injection and detectable tumor formation is long (R8 months). While this long latency period is consistent with the relatively indolent nature of lower-grade human gliomas, this time course should be considered in experimental designs using this model. Tumors are typically first detectable in their early stage on MRI scan prior to causing any neurological deficits in the mouse. However, neurological symptoms in the mouse may be an indicator of tumor growth, and thus mice should also be monitored periodically, particularly >4 months following injection. In addition, not all of the mice injected with AAV in this protocol will form gliomas. Some mice develop needle-track osteosarcoma tumors even when intracranial surgeries appeared to have been conducted optimally. To maximize accuracy of injection, we recommend performing practice surgeries on cadaver mice with trypan blue dye to confirm injection site (see troubleshooting 5). For increased tumor incidence and shorter latency, we recommend pursuing steps to generate allograft models as described above.

Potential solution
If stable positioning of the mouse is difficult to achieve, it is recommended that the person performing this procedure practice on cadavers. The mouse should be first secured into the nose fastener prior to securing with ear pins. The mouse's front teeth should fall within the bite hook (a small hole) in the nose fastener to facilitate stable positioning. The teeth should be securely inserted into the bite hook. This can be tested by gently pulling on the mouse's tail, at which point there should be a noticeable tension preventing the mouse's body from moving. If the ear pins do not stay within the mouse's ears, it may be helpful to position them in ears and tighten both screws together (as opposed to tightening one screw fully and then the other). The ear pins should be deep enough to allow for immobilization of the mouse, but not so deep as to cause physical trauma to the mouse.

Problem 2
Difficulty with drilling the skull (step 14).

Potential solution
Firm pressure should be applied with the handheld drill to pierce the skull. Pressure should be quickly released as soon as the skull has been successfully penetrated. Applying too much pressure will cause bleeding and should be avoided.

Problem 3
Air bubbles when drawing up AAV virus (step 15).

Potential solution
If air bubbles are present within the syringe, carefully expel the virus back into the tube and draw up again. If bubbles persist, remove the plunger from the syringe and back fill the syringe with R5 mL sterile PBS. Visible drops of PBS should fall from the needle tip upon backfilling. If drops do not appear, it may indicate a clog in the needle or loose connection of the needle to the syringe.

Problem 4
Lack of tumor growth.

Potential solution
Ensure injection site coordinates are accurate (1 mm posterior, 1 mm lateral to the bregma, 2.1 mm depth). See intracranial injection, step 13. Even with seeming optimal intracranial injections, some injected mice may not form tumors.

Problem 5
Skull-based tumor forms.

Potential solution
Needle-track tumors may form on the skull. This may be visible as a firm mass below the skin surface on the mouse's head that develops several months after injection. Ensure that depth coordinates are accurately zeroed on stereotactic frame at time of injection. Inject virus slowly to decrease leakage out of the brain. Inject low volume ($1 mL) virus to decrease leakage out of the brain.

RESOURCE AVAILABILITY
Lead contact Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Samuel McBrayer (samuel.mcbrayer@utsouthwestern.edu).

Materials availability
Reagents, resources, services, and equipment are outlined in the key resources table. Unique resources used in this protocol may be requested by contacting the lead contact, Samuel McBrayer (samuel.mcbrayer@utsouthwestern.edu). pAAV2-sgTrp53-sgAtrx-EFS-Cre AAV vector is available from Addgene (Plasmid #189977).

Data and code availability
This study did not generate databases nor code.